Many applications in the fields of pharmaceuticals, food and beverage manufacturing, clinical diagnostics, and life science research require that the concentration of a particular target analyte be determined accurately and precisely. Target analytes are often contained in very complex mixtures with many other species of similar molecules. Common samples containing target analytes include blood, cell lysates, cell culture media, process flowstreams, etc.
A highly selective assay is required for accurate and precise quantitation of analytes in such complex samples. Common assays used for this type of analysis are immunoassays. In immunoassays, antibodies are used to bind a target and immobilize it on a solid phase surface. This binding enables the target to be separated from the other molecules in the liquid sample by washing away the non-bound molecules. Following this separation, a second antibody to the target labeled with a marker such as an enzyme, fluorescent dye, etc. is used to produce a signal to sensitively indicate the amount of target bound to the solid phase. In some versions the target analyte itself may be labeled to produce a detectable signal. Immunoassays are selective, i.e., capable of discriminating between the target and other molecule species in the sample. They are also highly sensitive, i.e., capable of measuring concentrations in the ng/mL range or below.
Immunoassays, however, suffer from serious drawbacks for some applications. Immunoassays are relatively complex, with complicated and expensive reagent requirements. They can be challenging to automate. In a number of fields such as the manufacture of biopharmaceuticals, there is a great need for high precision in manufacturing applications to track yield or mass balance in the manufacturing process. The target analyte may be the protein product itself in a process flowstream. The product concentrations in such manufacturing applications can typically be in the μg/mL to mg/mL range. Thus, the high sensitivity of the immunoassay (typically in the ng/mL range or lower) can be an issue. A dilution of several orders of magnitude is often needed to bring the sample concentration into the range of the assay. This adds substantially to the complexity of the assay, and makes it difficult to achieve the required precision or reproducibility.
One common approach for this type of application is to use a high performance chromatography (HPLC) column packed with a resin that binds the target analyte in a highly selective fashion. Typical resins include an immobilized affinity ligand that selectively binds the target analyte. Alternatively, a more conventional chromatographic resin (such as ion exchange or reversed phase) may be used to selectively separate the target. After injecting a sample into an HPLC column, the column is washed, and the analyte is eluted with an agent that releases the analyte from the immobilized ligand. The analyte may be quantified using ultraviolet (UV) absorbance in the HPLC detector. An example of this process is the use of a protein A HPLC column to measure therapeutic monoclonal antibody (IgG) concentrations in samples from a cell culture production system. Such a test is highly selective for the target (IgG in this case), operates in the μg-mg/mL range with no sample dilution, and has very high precision (usually <5% coefficient of variation (CV)). However, affinity HPLC requires a complex and expensive instrumentation system. The system can run only one sample at a time, and each sample run typically takes 5-15 minutes. Thus, throughput on this system is limited to 4-12 samples per hour.
One way to improve sample throughput is to run multiple columns and samples at a time. Large-scale multiplexing with conventional HPLC systems is difficult and expensive because separate pumps, injectors, columns, and detectors are required for each sample channel or column. However, a number of relatively high-throughput devices and methods have been developed for sample preparation through solid phase extraction that could be applied to this type of quantitative assay.
One such device is the pipet tip column, which uses an air displacement pipet to move liquid in and out through a disposable tip packed with a selective binding resin. Multi-channel pipet systems (up to 96 or even 384 channels) can be used for high throughput applications. However, pipet tip columns have several disadvantages. The high-throughput platform requires an expensive automated robotic system. In addition, the flow of liquid through the packed bed in a pipet tip column is limited to a single port of entry and exit at the distal end of the tip. It is therefore very difficult to get quantitative washing and elution from the packed bed due to the mixing that occurs in both the pipet tip itself and the well containing the sample or wash buffer. Finally, the air-displacement method of moving the liquid makes it very difficult to control the flow rate of the sample, particularly at the relatively low flow rates required to obtain quantitative binding. Thus, while useful for qualitative sample preparation, pipet tip devices have proven to be unusable for quantitative analysis of targeted analytes.
An alternative type of high-throughput purification device uses a vacuum manifold to move liquid in a downward direction through multiple packed beds. Such devices are commonly used for solid phase extraction. As with the pipet tip columns, control of the flow rate is difficult, especially at the low flow rates required for quantitative extraction. Also, such manifolds must have columns present in all available vacuum positions or resulting vacuum leaks will render the device ineffective. This makes it difficult to modify the number of samples being run in a batch.
The use of gravity to draw the liquid downward through the packed bed is another purification method. For many applications, this is impractically slow. To speed the process, the columns may be placed in a centrifuge to increase the effective g-field to a level that provides a desired flow rate. Such “spin columns” are widely used for extracting a target from a sample in a variety of applications because they do not require complex instrumentation or equipment beyond widely available laboratory centrifuges. Virtually all commercially available spin columns are designed to work with common microcentrifuge tubes, which have an inner diameter (ID) of 9 mm and a volume of 1.5 or 2.0 mL. The outermost diameter of these devices must be significantly larger than 9 mm in order to prevent the resin-containing device from being driven into the tube. The resin bed volumes of these devices are typically 100 μL or greater.
For high-throughput analysis and convenient handling of large numbers of relatively small samples, it is highly desirable to be able to work within the Society for Biomolecular Sciences/American National Standards Institute (SBS/ANSI) microplate format standard. Many different types of standard microplates are available at low cost as molded plastic parts. Liquid handling devices, such as pipets, are also designed to work within the standard. The conventional SBS/ANSI 96-well microplates have wells spaced 9 mm on center. They are commonly designed for quickly reading the optical absorbance of all the wells using widely available optical plate readers.
Conventional spin column devices have several drawbacks. Currently available devices designed to work individually with microcentrifuge tubes are physically too large to function with microplates, both in terms of fitting into the 9 mm spacing and having a bed volume appropriate for the volume of the microplate wells. A number of multi-column spin column devices have been built that are in the form of microplates with a column in each well. If the user has 96 samples to run, these spin column plates work well. However, if fewer than 96 samples are to be run, the spin column plate is discarded after a first run, and the unused wells are wasted. Alternatively, the unused wells on the spin column plate are used in a second run, but this runs the risk of cross contamination. It would be advantageous to consume only one spin column per sample but to be able to collect the eluates in the wells of a conventional microplate for analysis.
Another drawback of conventional spin column devices is that they are susceptible to air entrainment. Air entrained within the spin column can have several deleterious consequences. These deleterious consequences include reduced flow rate which gives rise to flow rate variability between cartridges at a given “×g” force, as well as loss of binding capacity due to air blocking part of the packed bed. Both of these issues lead to greatly increased variability in the analytical results, and, in extreme cases, to air-locking of the cartridge which prevents the sample from being loaded at all. Trapped air bubbles, if large enough, can also reduce or block flow through the bed during centrifugation, particularly at the low g-fields required for quantitative binding. There is a need for spin column devices that are not susceptible to air entrainment.
Yet another drawback of conventional spin column devices is that they are susceptible to drying. Some types of solid phase supports spin column devices must be stored and shipped in a wet condition. These supports include common gel-type chromatography media such as agarose, dextran, cellulose, and various synthetic polymer hydrogel materials, such as poly(methacrylates). These materials can suffer irreversible collapse of the gel structure on drying, which significantly degrades performance. There is a need for spin column devices that are less susceptible to drying.